Endothelial cell CD36 regulates membrane ceramide formation, exosome fatty acid transfer and circulating fatty acid levels

Endothelial cell (EC) CD36 controls tissue fatty acid (FA) uptake. Here we examine how ECs transfer FAs. FA interaction with apical membrane CD36 induces Src phosphorylation of caveolin-1 tyrosine-14 (Cav-1Y14) and ceramide generation in caveolae. Ensuing fission of caveolae yields vesicles containing FAs, CD36 and ceramide that are secreted basolaterally as small (80–100 nm) exosome-like extracellular vesicles (sEVs). We visualize in transwells EC transfer of FAs in sEVs to underlying myotubes. In mice with EC-expression of the exosome marker emeraldGFP-CD63, muscle fibers accumulate circulating FAs in emGFP-labeled puncta. The FA-sEV pathway is mapped through its suppression by CD36 depletion, blocking actin-remodeling, Src inhibition, Cav-1Y14 mutation, and neutral sphingomyelinase 2 inhibition. Suppression of sEV formation in mice reduces muscle FA uptake, raises circulating FAs, which remain in blood vessels, and lowers glucose, mimicking prominent Cd36−/− mice phenotypes. The findings show that FA uptake influences membrane ceramide, endocytosis, and EC communication with parenchymal cells.

Long-chain fatty acids (FAs) play a prominent role in membrane structure and function. They are important in systemic energy metabolism, with mitochondrial FA oxidation serving as an efficient ATP source for heart and muscle, while FA esterification into triglycerides confers energy storage capacity to adipose and other tissues. Fatty acid oxidation also influences health of stem cells and tissue renewal [1][2][3] . Fluctuations in cellular FA uptake and utilization during the physiological transitions in energy metabolism are critical for maintaining homeostasis as the circulating FAs are routed to specific tissues 4,5 . During fasting or exercise when glucose supply is reduced, adipose tissue lipases mobilize FAs from stored triglycerides (TGs) to increase supply of circulating FAs. Simultaneously, heart and muscle tissues upregulate FA uptake and oxidation while reducing glucose utilization thus sparing glucose reserves. In contrast, as feeding makes more glucose available, its usage by heart and muscle rises and more FAs are directed for storage in adipose depots 5,6 .
In tissues, FA uptake occurs mainly at the level of capillaries and begins with FA transfer across microvascular endothelial cells (MECs). Surface high-affinity FA receptors on MECs are required to recognize FAs and work in concert with intracellular FA-binding proteins and FA acyl-CoA ligases to promote rapid lipid accumulation [7][8][9] . In the circulation, FAs are bound to albumin or esterified in lipoprotein triglycerides. FA release from lipoproteins is mediated by intravascular lipoprotein lipase (LpL) and its endothelial cell (EC) anchor protein Gpihbp1. The level of unesterified FA released by LpL at the EC surface is likely to be higher than that of FA dissociated from albumin in the circulation 10 but is unknown 11 .
Several proteins influence cellular FA uptake including members of the FA transport (FATP) family, the FA binding proteins (FABPs), the acyl-CoA synthases, and cluster of differentiation 36 (CD36). These proteins can function at different levels of the uptake process involving plasma membrane transfer or intracellular channeling of the FA, as reviewed 12,13 . The plasma membrane glycoprotein CD36 was identified as a FA transporter through affinity labeling with a reactive membrane-impermeable sulfosuccinimidyl derivative of oleic acid, SSO 14 , and its physiological role in tissue FA uptake was demonstrated in rodents and humans 15,16 . In addition to albumin bound circulating FAs, CD36 transfers FAs released by LpL from very low-density lipoproteins (VLDL) 17 , however, FA uptake from chylomicron hydrolysis is CD36-independent 18,19 . Crystal structure of CD36 and family members which includes SR-B1 and Limp2 uncovered a common internal lipid transport tunnel 20 and long-chain FAs were identified in the tunnel of crystal CD36 21 . The FAs bind in a hydrophobic surface pocket of CD36 22 that connects to the tunnel based on modeling the crystal structure 23 .
CD36 is highly expressed in the capillary endothelium, where it is part of the gene expression signature that defines the lipid handling of microvascular cells 9,24,25 . Strong evidence derived from EC-specific deletion of various transcription factors supports the gatekeeping function of the endothelium in tissue FA uptake 7,24,26 , and EC-specific deletion of CD36 supports its critical role in the process 9 . Mice lacking CD36 in endothelial cells (EC-Cd36 −/− ) have impaired FA uptake into adipose tissue, heart, skeletal muscle 9 , and the gastric mucosa 3 . EC-Cd36 −/− mice also recapitulate the higher plasma FAs and enhanced glucose disposal observed in germline Cd36 −/− mice 9 .
In this study, we examined the mechanism by which ECs transfer exogenous FA to parenchymal cells. We document that FAs elicit CD36-dependent phosphorylation of caveolin-1 (Cav-1) and generation of ceramides in caveolae, which associate with caveolae budding into internal vesicles (IVs) containing FA, CD36, Cav-1, and ceramides. These vesicles are subsequently released at the basolateral membrane as exosome-like small extracellular vesicles (sEVs) that deliver FAs to tissue cells. We define the FA-sEV pathway biochemically using various approaches and document it's in vivo contribution using a reporter mouse with EC expression of fluorescent CD63. Inhibiting the sEV pathway in mice reduces FA uptake by muscle fibers and increases circulating FA levels.

Results
Fatty acids are endocytosed apically by endothelial cells (ECs) and secreted basolaterally in exosome-like small extracellular vesicles (sEVs) We previously reported on critical contribution to tissue FA uptake of EC CD36 9 . CD36 deficient mice show reductions in FA uptake by heart, muscle, fat, and gastric tissues 3,27 that are replicated by specific deletion of EC CD36 3,9 . Microvascular ECs in heart, muscle, and lung where tissue FA uptake occurs have high expression of CD36 28 , while levels are lower in macrovascular ECs 25 . To investigate the pathway for FA uptake in microvascular ECs (MECs) we used mouse primary ECs (mMECs) isolated from lungs of wild-type or CD36 −/− mice 9,29 , and human-derived dermal microvascular ECs (hMECs) [30][31][32] . These capillary-derived cell types have high expression of CD36 9,24,25 . To trace the FA, we used biologically relevant alkyne-modified analogs of longchain FAs, amenable to click chemistry that can be reacted post cell fixation with azide-fluorochromes 33,34 (Fig. 1a). The reactive tag in alkyne-modified FAs is small, consisting of only two carbon atoms connected by a triple bond and results in minimal disruption of cellular FA handling [35][36][37] . Microvascular ECs were exposed to 15 μM alkyne FAs; oleic or palmitic acids (OA, PA, respectively). Cells were then fixed, the alkyne FAs were conjugated to Alexa Fluor 555 azide (red) followed by immunostaining for CD36 (green). Fatty acid uptake studies traditionally add FAs complexed with bovine serum albumin (BSA) to solubilize the FA and relatively high FA concentrations (>100 µM) are used to keep FA dissociation from becoming limiting 38 . In the present work we used mostly short incubations (10-15 min) and the alkyne FA concentrations found optimal for our microscopic studies were low (10-25 µM), so BSA was not required. We compared uptake of FA complexed to 0.2% BSA to FA without BSA. Omission of BSA did not significantly alter the CD36/FA puncta (Fig. 1b, c). Quantification of CD36-FA co-localization 39 yielded a strong Pearson's coefficient of 0.57-0.65 for OA and PA, which was little changed whether BSA was included or not during uptake, as determined by two-way ANOVA adjusted for multiple comparisons (Fig. 1d, p > 0.05, n = 3 experiments, cells analyzed each experiment = 15-25). Thus, we omitted BSA from our microscopy studies to avoid the factors introduced by BSA FAbinding and BSA transcytosis. However, BSA was kept in the medium during long incubations when higher FA concentrations were needed, as indicated.
Caveolin-1 (Cav-1) was reported to bind FA in adipocytes 40 . Immunostaining for Cav-1 (cyan) in MECs was robust, consistent with its known abundance in microvascular ECs 41 . Cav-1 appeared to dot the CD36-OA puncta ( Fig. 1e) but in contrast to the tight co-localization of CD36 and OA, co-localization of Cav-1 and OA was low and had a Pearson's coefficient of 0.26 (Fig. 1d, n = 3 experiments, cells analyzed per experiment = 15). In addition, only some of immunostained Cav-1 (cyan) associated with the OA/CD36 complex (Fig. 1e, insets) consistent with the more ubiquitous role of Cav-1 in EC biology 42 .
Uptake of alkyne OA, quantified by its mean fluorescence intensity, was markedly reduced by CD36 deletion in mMECs of Cd36 −/− mice as compared to mMECs from wild-type (WT) mice (Fig. 1f). Inhibition of CD36 by SSO, which binds the protein irreversibly 22,43 substantially reduced hMEC FA uptake, measured in real-time using Bodipy-C12 FA, a FA analog that competes well with native FAs 44,45 (Fig. 1g). The uptake slope was reduced by 70% (0.116 to 0.032) and the area under the curve (AUC) by 65% (Fig. 1h, p < 0.0001, n = 4 experiments). Together the data in Fig. 1 show that CD36 in MECs recognizes alkyne FA and internalizes the FA with CD36 and Cav-1. Results with both alkyne OA and Bodipy-C12 FA showed CD36 is required for optimal FA uptake by MECs in line with previous findings in mice where deletion of CD36 in ECs reduced tissue FA uptake 9 .

Role of caveolae in FA uptake by MECs
Cav-1 is integral to the formation and maintenance of caveolae through its interaction with various proteins and lipids 42,46 . The caveolae are dynamic 50-100 nm bulb-shaped plasma membrane (PM) invaginations 47 abundant in MECs. Caveolae function in transcytosis of macromolecules from the luminal/apical side to the basolateral subendothelial space 48,49 . In adipocytes, caveolae were linked to FA uptake 50,51 and FAs were reported to induce caveolae endocytosis in these cells 50 . Using immune electron microscopy (EM) we monitored Cav-1 trafficking as a marker of caveolae movement in hMECs treated with OA for 5, 15, or 60 min. Addition of OA visibly enhanced Cav-1 internalization at the apical membrane, and Cav-1 movement to the basolateral side was consistent with OA inducing Cav-1 transcytosis (Fig. 2a).
The data showing FA induced transcytosis of Cav-1 and the inhibitory effect on FA uptake of the Cav-1 Y14F mutant in MECs treated with FA led us to further examine how FA regulate caveolae dynamics using EM. Addition of OA increased the number of caveolae at the apical EC surface or just endocytosed underneath the surface (Fig. 3a-c, quantified in graph as caveolae/unit area, p < 0.05, 10-13 cells analyzed per group). When we exposed the cells to nanogold PA we could identify PA in caveolae that are being internalized or that already dissociated forming intracellular vesicles (IVs) (Fig. 3d). The PA was also detected in multivesicular late endosomes also referred to as multivesicular bodies (MVBs) (Fig. 3e, and Supplementary Fig. 1a, b). The MVBs are distinguished by their content of intraluminal vesicles formed by   inward budding of the endosome membrane. Vesicles in MVBs can either be degraded by MVB fusion with lysosomes or are released from the cell as small extracellular vesicles (sEV) by MVB fusion with the plasma membrane. The latter outcome was observed as vesicles containing the nanogold PA were released at the EC basolateral side, and these vesicles measured less than 100 nm in diameter consistent with their caveolae origin ( Supplementary Fig. 1a, b). To confirm that the internalized FAs are released in sEVs, we isolated sEVs from media of ECs incubated with nanogold PA. Electron microscopy detected PA in sEVs (Fig. 3f) which were also positive for CD36 and Cav-1 using nanogold immunostaining (Fig. 3g).

Characterization of sEVs
The sEVs secreted by hMECs were isolated and subjected to analysis following the guidelines of the International Society for Extracellular Vesicles, ISEV 58 . The sEVs had an average diameter of about 100 nm (Fig. 3h) and expressed the common sEV (exosome) markers CD9, CD81 and excluded ER calnexin (CANX) and lysosomal Rab7, a marker of late endosome/lysosome and less frequent exosomal marker ( Fig. 3i). Size distribution of sEVs from mMECs was broadly similar and did not differ whether cells were or were not treated with OA and if they expressed or did not express CD36 (Fig. 3j). The sEVs isolated from WT or CD36 −/− MECs all expressed exosome markers, CD81, and TSG101 related to MVB biogenesis, and excluded CANX (Fig. 3k). No differences were observed in average sEV size (80-90 nm) between WT and CD36 −/− mMECs and whether cells were or not OA-treated (Fig. 3L, n = 12 preparations).
Visual evidence for presence of CD36, Cav-1, and OA in sEVs was obtained by confocal microscopy of hMECs grown on transwell filters, which allows imaging and visualization of events at the basolateral side of the polarized ECs. In hMECs treated with alkyne OA, threedimensional rendering of confocal stack images showed rosette-like clusters of vesicles with OA, CD36, and Cav-1 (Fig. 4a) that bud from the basolateral EC side (Fig. 4b), consistent with MVBs fusing with the membrane to release sEVs. In addition, sEVs that are positive for CD36 (green), oleic acid (red), and Cav-1 (magenta) could be observed at the cell periphery and in the adjoining extracellular space (Fig. 4c, d, white arrows). We also immunostained for CD63, a common sEV marker and identified sEVs positive for CD36, CD63, and OA that were released by hMECs in the extracellular space ( Supplementary Fig. 1c, d, white arrows).

CD36 regulation of membrane ceramide mediates FA-sEV secretion
Vesicles generated by endocytosis are either routed to intracellular organelles such as lysosomes or targeted for release into the extracellular space. Ceramides generated from sphingomyelins by neutral sphingomyelinases are important for sorting vesicles into the lumen of MVB for secretion 59 . We immunostained for ceramides 60 to determine if ceramide generation is involved in OA transcytosis. Antibody specificity was validated by SMase treatment, which enhanced ceramide staining, while staining was eliminated by co-treatment with ceramidase which hydrolyzes ceramides to sphingosine (Supplementary Fig. 3a, b).
The sorting of specific shape-inducing lipids between caveolae and the bulk membrane regulates caveolae formation, stability, and dynamics 61 . We performed targeted lipidomics on the sEVs to examine if OA addition, which induces caveolae endocytosis, associates with noticeable changes in the shape-inducing ceramides and lysophosphatidylcholine (LPC) known to impact caveolae stability 61 . Among the three ceramide species identified in sEVs, we measured Cer 18:1;O2/24:1 previously reported in the few studies that measured sEV ceramides 62  Ceramide formation induced by OA addition was observed at cell membrane caveolae invaginations as well as in intracellular vesicles ( Fig. 5e, upper panel). Plasma membrane neutral sphingomyelinase 2 (nSMase2) located in the internal leaflet of caveolae 63 hydrolyzes sphingomyelins to generate ceramide ( Fig. 5e cartoon, lower panel) and has been implicated in exosome formation. GW4869 a specific inhibitor of nSMase2 has been widely used to demonstrate nSMase2 involvement in various functions [64][65][66] . GW4869 prevents enzyme activation by plasma membrane anionic phospholipids such as phosphatidylserine 67,68 and inhibits exosome secretion 59,69-72 . We found that GW4869 (Fig. 5f) effectively inhibited FA-induced ceramide formation and blocked FA internalization linking ceramide generation by nSMase2 to caveolae endocytosis and FA uptake (Fig. 5f). To further validate these findings, we tested effect of GW4869 on real-time FA uptake. Uptake was significantly reduced in GW4869-treated cells as compared to vehicle-treated controls (Fig. 5g, p < 0.0001, n = 4 experiments).
The above data demonstrate that both depletion of CD36 and inhibition of nSMase2 block ceramide generation and FA internalization. We showed earlier in Fig. 2, that OA induces Cav-1 phosphorylation at Y14, a Src kinase site, and this phosphorylation is known to disrupt the caveolae coat promoting caveolae scission from the membrane 53 . We examined the role of Src as CD36 was reported to associate with Src-related protein kinases in MECs 54 . We confirmed the CD36-Src interaction through immunoprecipitation (Fig. 6a). Src inhibition by Src inhibitor-1 blocked OA ability to induce phosphorylation of Cav-1 Y14 (Fig. 6b, c, p < 0.0001, n = 3 experiments) and it reduced FA uptake by hMECs measured in real time (Fig. 6d, p < 0.0001, n = 4 experiments). Furthermore, confocal images of mMECs and hMECs (Fig. 6e, f) pretreated with Src inhibitor-1 prior to adding OA, suppressed FA internalization. The OA and the generated ceramide remained at the cell periphery with Cav-1. Together the data shown in Figs. 1-6 support a molecular mechanism of FA uptake that involves FA interaction with CD36 inducing Src phosphorylation of Cav-1 Y14 which disrupts the Cav-1 coat and consequently Cav-1 interaction with other caveolae proteins leading to caveolae endocytosis 42,73 . Src did not eliminate ceramide formation in the membrane suggesting that its effect on endocytosis is at least in part independent of nSMase2 activation.
The EC caveolae transcytosis pathway for FAs, was supported by RNASeq data collected in hMECs without and with CD36 KD which showed altered expression of genes related to regulation of caveolae endocytosis (EHD3, Dynamin), vesicular traffic and sorting (VPS37B, RAB36) and vesicle exocytosis (RAB3C) (Supplementary Tables 1  and 2).

Fatty acid uptake requires FA remodeling of cellular actin
Since endocytosis and vesicular traffic depend on the cytoskeleton 74 , we next examined if rearrangement of cellular actin a hMECs, treated with either control siRNA or CD36-specific siRNA, were serum-starved (4 h) and supplemented with alkyne OA (15 µM, 10 min). After the click reaction, cells were immunostained for CD36 and ceramide using secondary antibodies conjugated to Alexa Fluor 647 and 488, respectively. Lower panel, CD36 knockdown (KD) suppresses OA uptake and ceramide formation in hMECs. Adjoining graph, Ceramide mean intensity in control and CD36KD cells. n = 3 independent preparations, number of cells counted per preparation: 15-25, ****p < 0.0001 by unpaired t test. Data are means +/− SEM. Scale bars: 10 µm. b Alkyne PA treated control or CD36KD hMECs, were immunostained for ceramide. Co-localization of alkyne PA with ceramides is shown. Adjacent graph shows ceramide mean intensity in control and CD36KD cells. n = 3 independent experiments, *p < 0.05 by unpaired t test number of cells counted/experiment 10-25. Data are means +/− SEM. Scale bars: 10 µm. c, d High-performance LC/MS lipidomics analysis on sEVs isolated from mMEC treated with 200 µM OA. c Ceramide Cer 18:1;O2/24:1 content in sEVs from WT and CD36 −/− cells was similar. d Ceramide Cer 18:1;O2/24:1 content increased with oleic acid treatment in sEVs from CD36 +/+ cells and decreased in sEV from CD36 −/− cells, n = 6 independent preparations, **p < 0.01 by unpaired t test. Data are mean values +/− SEM. e Ceramide generation in caveolae. Co-localization of ceramide and Cav-1 in OA-treated hMECs: Cav-1 and ceramide are shown to colocalize in plasma membrane caveolae (arrows) and in intracellular vesicles. Scale bar 10 µm. Lower panel in (e) shows a cartoon summary of plasma membrane sphingomyelinceramide turnover: Neutral sphingomyelinase 2 (nSMase2) localizes to the inner leaflet of caveolae and upon activation generates ceramide, which can be hydrolyzed by ceramidase to sphingosine. Scale bar: 10 µm. f The nSMase2 inhibitor GW4869 reduces ceramide formation and OA uptake: hMECs were pre-incubated 10 min with vehicle (DMSO) or GW4869 (10 µM) before adding alkyne OA (15 µM, 10 min). Scale bars: 10 µm. g Ceramide formation is critical for FA uptake. Real-time FA uptake (AUC) in hMECs treated with either vehicle or GW4869. n = 4 independent experiments, ****p < 0.0001 by unpaired t test. Data are means +/− SEM.
Article https://doi.org/10.1038/s41467-023-39752-3 is required for OA uptake by MECs. OA addition was found to rapidly alter morphology of actin stress fibers in hMECs causing peripheral fiber clustering close to actin-containing cellular extensions (Fig. 7a, arrows). The extensions, which protruded into the extracellular space between cells, appeared to be decorated with sEVs (Fig. 7b, insets). Actin-containing extensions can help transfer sEVs to recipient cells 75,76 . Actin remodeling in response to OA was demonstrated by a change in ratio of filamentous (F) to monomeric globular (G) actin (Fig. 7c, p *< 0.05, n = 3 experiments) and by increased CD36 association with actin and cofilin, a key protein that depolymerizes actin and maintains a dynamic F/G actin equilibrium (Fig. 7d). In addition, OA induced ezrin phosphorylation at activatory threonine 567 while it induced dephosphorylation of cofilin at inhibitory serine 3 (Fig. 7e), both changes would facilitate actin remodeling. To directly test requirement of actin remodeling in FA uptake we examined the effect of latrunculin B, which binds actin monomers preventing them from polymerizing thereby affecting G-actin recycling 77,78 . Pretreatment of hMECs with latrunculin resulted in substantial reduction of FA uptake (Fig. 7f, p < 0.0001, n = 4 experiments). In contrast, FA uptake was only slightly suppressed by Y-27632 which inhibits the Rho GTPase effectors ROCK1 and 2 (Rho-associated coiled-coil-containing protein kinase) that regulate actin organization for cell motility and other cellular functions 79 (Fig. 7g, p < 0.05, n = 4 ctrl, 3 Y-27632). In line with relevance of actin remodeling in facilitation of CD36-mediated FA uptake, CD36 knockdown in hMECs altered expression of proteins implicated in actin organization such as fascin, profilin, myosin light chain kinase (MLCK), coronin7 and fibrillin 2 (Supplementary  Tables 1 and 2).

Endothelial cells rapidly transfer FA-sEVs to myotubes
Transfer of sEVs from MECs for uptake by myotubes was visualized in a polarized setting with hMECs cultured in the upper chamber of a transwell, and myotubes placed underneath in the lower chamber (Fig. 8a). Barrier integrity of the MEC monolayer was assessed by transendothelial electric resistance (TEER), which measures monolayer impermeability based on resistance to ionic flux between cells (paracellular) 80 . As the MECs seeded on the filter increased in number, and intercellular junctions started closing the gaps between cells, electric resistance increased, reflecting the progressive decrease in monolayer permeability, as is shown in Fig. 8b left panel. TEER reached a plateau (maximum resistance) when the barrier was fully formed. The TEER plateau was not altered by OA addition indicating that OA did not breach the barrier (Fig. 8b right panel). Further confirmation of barrier integrity used C12 Bodipy FA (red) and Dextran-FITC (green) added to the upper chamber in absence or presence of OA. Fluorescence of both was monitored in the lower chamber. Dextran transfer shown as percent of input in the top compartment was very low, around 0.02% at 30 min which is our assay time, and it remained low (0.2% of input) at 240 min indicating stable monolayer integrity over time. Transfer of both dextran and Bodipy was unaffected by added OA (Supplementary Fig. 4a).
The hMECs were transduced to express the red fluorescent CD63 (RFP-CD63), a plasma membrane protein and sEV marker, before culture on the transwell filters until 3-5 days post confluence. There was low transfer of CD63 to myotubes during a 30 min basal period as deduced from the diffuse red signal in Supplementary Fig. 4b. OA addition to hMECs in the upper chamber resulted in detectable red CD63 transfer from hMECs to myotubes reflecting sEV uptake ( Supplementary Fig. 4b, middle and bottom panels) and the red CD63 signal merged with green CD36 immunostaining (Supplementary Fig. 4b, middle and bottom panels) consistent with transfer of EC CD36 in CD63 sEVs.
We confirmed in transwells that the sEV transfer mechanism applies to PA and OA by adding alkyne OA or PA to hMECs on filters in the top chamber, then evaluating transfer to myotubes in the bottom well. Red signal from OA (Fig. 8c, upper panels, middle lane) and PA (Fig. 8c, lower panels, middle lane) was measured in myotubes and co-localized with CD36 ( Fig. 8c middle  lanes, merged images). Importantly, this transfer was blunted, becoming almost undetectable when the hMECs were treated with GW4869, a neutral sphingomyelinase 2 (nSMase2) inhibitor, which blocks sEV generation. The inhibition observed was comparable for OA and PA (Fig. 8c adjacent graphs, n = 3, p < 0.01 for OA and p < 0.001 for PA). These results showed that the FA (OA or PA) transfers to myotubes in sEVs and blocking sEV generation markedly suppresses the transfer. These results also indicate that the major FA transfer pathway is across EC through transcytosis and sEV generation, as the monolayer is impermeable to paracellular FA flux. We next visualized OA transfer from hMECs to myotubes in labeled sEVs. We performed the transwell studies with alkyne OA added to upper chamber hMECs expressing the RFP labeled exosome marker CD63. The OA recovered in lower chamber myotubes was clicked with fluor-azide as before. In this experiment, unlike in Fig. 8c, OA was visualized with Alexa fluor-488 (green) instead of fluor-555 to distinguish it from the red RFP-CD63. During a 30 min basal period (no OA added) little RFP-CD63 transfer to myotubes was seen (Fig. 8d, top panel). OA addition (Fig. 8d, middle and bottom panels) stimulated transfer of RFP-CD63 (red) and OA (green) to myotubes, where OA and CD63 co-localized forming intracellular yellow puncta, as highlighted in the two insets of the middle panel and in the inset of the bottom panel. These data supported FA transfer from hMECs to myotubes in RFP-CD63 sEVs. Taken together, these data show that FAs are delivered by ECs to parenchymal cells through generation of sEVs.
The sEV pathway contributes to tissue FA uptake and regulation of blood FAs We next examined in vivo operation of the sEV-exosome pathway in FA uptake. We used a reporter mouse with EC-specific expression of emeraldGFP-tagged CD63 (emGFP-CD63) 84 to visualize transfer of FAexosomes from ECs to muscle fibers. Alkyne OA was given retroorbitally to the mouse and 30 min later, hindlimb plantar flexor muscles were dissected. Frozen sections (10 µm) were fixed, processed for FA click reaction, and immunostained for CD36. Confocal images of muscle fibers (Fig. 9a, top panel) showed expression of CD36 (white), EC-emGFP-CD63 (green), and OA (red) at the periphery of muscle c hMEC transfer OA and PA to myotubes through sEVs. Human myoblasts were grown on coverslips and differentiated into myotubes. The hMECs were grown on filters in transwells until 3-4 days post confluence. The myotubes on coverslips were placed in the bottom wells of the transwells. c Alkyne OA (upper panels) or PA (lower panels) was added (25 µM, 30 min) in presence or absence of the nSMase2 inhibitor GW4869 (GW, 10 µM, 10 min). Myotubes were clicked and processed for CD36 immunostaining and confocal microscopy. The percent GW4869 inhibition of OA and PA flux to myotubes is shown in adjacent graphs. n = 3 independent experiments, **p < 0.01 for OA, ***p < 0.001 for PA by unpaired t test. Data are means +/− SEM. Scale bar: 10 µm. d Alkyne OA transfers with the exosome marker CD63 when hMECs express RFP-CD63 (red). Alkyne OA was added to the hMECs and 30 min later myotubes were processed to visualize OA and CD63. Top control panel: Myotubes under control hMEC (no OA added) showed low diffuse red CD63 signal. Panels + OA: Myotubes cultured below OA-treated hMECs showed bright intracellular red CD63 puncta that were also positive for OA (green azide Alexa Fluor-488). Note green OA signal was used to distinguish it from the red RFP-CD63 In the middle +OA panel, arrows point to puncta positive for CD63 and OA. Inset: Myotubes at higher magnification showing OA/CD63 vesicles inside and outside the cell (arrows) and an additional higher magnification highlights OA/ CD63 vesicles with perinuclear localization. Bottom panel shows an experiment similar to that in the middle panel to further illustrate the yellow merge of hMECtransferred OA (green) with CD63 (red) inside myotubes. Scale bars: 10 µm. e hMEC FA-sEVs alter gene expression in myotubes. qPCR analysis of myotubes treated 2 h with sEVs from ECs with and without OA treatment. CPT1a: carnitine palmitoyl transferase a, ACSL3: Acyl-CoA synthase long chain 3, FABP3: FA binding protein 3, PPARδ: Peroxisome proliferator activating receptor delta, eNOS: endothelial nitric oxide synthase, VEGFR2: Vascular endothelial growth factor receptor 2, IRS1: Insulin receptor substrate 1, Glut1: Glucose transporter 1, HK2: hexokinase 2. n = 3-6 independent preparations, *p < 0.05 by unpaired t test. Data are means +/− SEM.
fibers as well as on myofibrils inside muscle fibers which appeared as puncta inside the fibers. Previously, lipid droplets and mitochondria were shown to associate with the myofibrils of muscle fibers 85 . Highmagnification images to highlight the myofibrils (Fig. 9a, bottom panel) were used to quantify co-localization of OA and CD63 inside the fibers. This yielded a strong Pearson coefficient of 0.6 consistent with OA traveling in CD63 labeled sEVs to myotubes (Fig. 9A, adjoining graph n = 3 animals, 8-10 muscle fibers/animal). The co-localization coefficient of CD36 and emGFP-CD63 (0.43) was lower, likely reflecting immunostaining of both endogenous muscle CD36 and the CD36 transferred from ECs.
We tested importance of the sEV pathway in muscle uptake of circulating FAs by giving wild-type mice the nSMase2 inhibitor GW4869 (intraperitoneal, 4 days) and then a retro-orbital injection of alkyne OA. Muscles were excised 30 min after the OA, sectioned, fixed, clicked, and stained as before. Confocal microscopy showed that GW4869 reduced by about 50% OA staining of muscle fibers (Fig. 9a, adjoining graph, p < 0.001, n = 3 animals per group, 10-15 muscle fibers/animal).
Muscle uptake of circulating OA was paralleled with OA depletion from blood vessels (Fig. 9c arrows). Muscle fibers of vehicle-treated WT mice given alkyne OA stained brightly for CD36 (green) and OA (red) while a nearby blood vessel expressing CD36 (Fig. 9c, top panel, arrows) had little red OA staining indicating OA had already transferred out of the vessel. The opposite was seen in muscle of GW4869-treated mice where fibers had low OA staining while a nearby blood vessel expressing CD36 retained substantial OA signal (Fig. 9c lower panel, and Supplementary Fig. 6a, lower panel).
To confirm that inhibition of exosome biogenesis in vivo causes endothelial retention of circulating FA, we measured plasma-free FA (NEFA) in mice given GW4869. We found (Fig. 9d) that these mice had increased blood FAs when tested fed or after a 5 h fast (Fed: vehicle vs GW4869 p = 0.0001; Fasted: vehicle vs GW4869 p = 0.0069, n = 4-5 mice, two-way ANOVA with adjustment for multiple comparisons). Interestingly, GW4869 reduced blood glucose in mice, consistent with the reduced muscle FA uptake (Fig. 9e, p < 0.05, n = 4-5 mice). The effects of GW4869 on blood FAs and glucose mimic effects observed in Cd36 −/− and EC-Cd36 −/− mice 9,86 . In line with this, GW4869 treatment did Adjoining graph shows Pearson's co-localization coefficient for CD63/OA and CD36/CD63 (n = 3 animals, 8-10 individual muscle fibers/ animal). b OA uptake by muscle fibers is suppressed by GW4869. Mice were injected with vehicle (DMSO) or GW4869 (2.5 µg/g intraperitoneally for 4 days) then given alkyne OA (25uM) retro-orbitally. GW4869 reduced OA uptake by muscle fibers as quantified in adjoining graph. n = 3 mice per group for vehicle and GW4869 treatment, 10-15 individual muscle fibers/mouse ***p < 0.001, unpaired t test. Data are means +/− SEM. Scale bar, 10 µm. c GW4869 induced reduction in OA uptake by muscle fibers associates with trapping of the OA in blood vessels. Skeletal muscle from a vehicle-treated mouse (upper panel) shows strong OA uptake into fibers and OA absence from a nearby CD36-expressing vessel (arrows). Muscle from a GW4869-treated mouse (lower panel) showed reduced uptake into fibers, and OA retention in a nearby CD36-expressing vessel (arrows) can be seen. Scale bar 20 µm. d GW4869 increases blood levels of non-esterified FAs (NEFAs). Mice were treated with GW4869 or vehicle, as in (c) at end of the dark period (fed) or after a 5 h fast. NEFAs were measured in tail vein blood. n = 4-5 mice/group, ****p < 0.0001, ***p = 0.0002 by two-way ANOVA adjusted for multiple comparisons. Data are means +/− SEM. e GW4869 reduces blood glucose. Mice (vehicle or GW4869 treated) were fasted for 5 h and blood glucose measured in tail vein blood. *p < 0.05 by unpaired t test, data representative of two cohorts n = 4-5 mice per/group. Data are means +/− SEM. Note: Blood glucose in C57bl6 mice (fed or fasted) are normally higher as compared to levels in other commonly used strains 109, 110 . not alter FA or glucose levels in Cd36 −/− mice ( Supplementary Fig. 6b). We also examined effect of in vivo inhibition of Src by Scr inhibitor-1 on alkyne PA uptake. Src-1 blocks Cav-1 Y14 phosphorylation and caveolae endocytosis, without eliminating ceramide formation (shown earlier in Fig. 6e, f). Animals injected with Src-1 showed reduced alkyne PA uptake in muscle fibers as compared to vehicle-treated mice. Src inhibition also associated with more retention of PA in blood vessels, similar to alkyne OA vessel retention in GW4869-treated mice (Supplementary Fig. 6c, d). Injection of Src-1 also significantly raised blood FAs ( Supplementary Fig. 6e, p < 0.01, n = 5-6 mice per group) Together these data support the physiological role of the sEV pathway in endothelial transfer of circulating FAs to underlying tissue cells.

Discussion
Since its identification as a FA transporter, numerous studies have documented role of CD36 in tissue FA uptake and utilization in mice and humans, as recently reviewed 12,13,87 . Endothelial cell-specific deletion of CD36 in mice showed EC CD36 is required for tissue FA uptake and recapitulated many metabolic effects of global CD36 deficiency 9 . This study documents that ECs transfer FAs from the circulation to underlying tissue through sEVs. Blocking sEV formation increases blood FA levels and replicates phenotypes found with CD36 deletion.
We described how FA interaction with CD36 in the apical membrane of ECs triggers CD36/Src coordinated FA internalization through caveolae budding into intracellular vesicles (IVs) that travel and are secreted as sEVs at the basolateral side. We used a combination of approaches including microscopy, biochemical analysis, and real-time FA uptake assays to identify the transcytosis pathway. We visualized FA internalization by CD36 and Cav-1 using confocal microscopy coupled with click chemistry of alkyne OA or PA. Using TEM with nanogold PA, we highlighted FA endocytosis into caveolae-derived intracellular vesicles (IVs), sorting of the FA-IVs into MVBs, and the secretion of sEVs that contain FA, CD36, and Cav-1. Secretion of these sEVs was also shown by confocal microscopy. The isolated sEVs have common exosome markers and a diameter between 80-100 nm consistent with their caveolae origin. Three critical mechanisms were identified in EC CD36-mediated transcytosis of FAs: First, FA-induced Src phosphorylation of residue Y14 in Cav-1. Both Src inhibition and the expression in MECs of Y14 mutated Cav-1 were shown to suppress FA uptake by confocal microscopy and real-time FA uptake. Second, FA-induced generation of ceramides by membrane sphingomyelinase. Inhibition of membrane nSMase2 suppressed FA-induced ceramide formation, FA internalization, and FA uptake by MECs. Third, FA-induced rearrangement of cellular actin. The FAs triggered phosphorylation of actin remodeling proteins cofilin and ezrin and enhanced CD36 association with cofilin and actin. Inhibition of actin remodeling reduced FA uptake. The above data documented the details of CD36 facilitated EC FA transcytosis and secretion in sEVs as highlighted in the schematic model of Fig. 10.
Both FA-treated and untreated ECs, and ECs with or without CD36 expression released exosome-like sEVs with a diameter between 80-100 nm. This is consistent with ECs releasing sEVs at a continuous low basal rate while addition of FAs rapidly upregulates caveolae endocytosis and sEV formation. Due to the basal output of sEVs and the FA-containing sEVs being a subset of the total sEVs secreted, our lipidomics measurements represent the total sEV pool. However, despite this limitation, conclusions can be made related to the changes induced by OA treatment in WT versus CD36 −/− cells. The percent OA in sEVs doubled with FA treatment when ECs expressed CD36 but not when ECs lacked CD36. A couple of observations suggested that FA addition disrupts distribution of specific, shape-inducing lipids, known to impact caveolae stability 61 . We measured opposite OA-induced changes in sEV ceramide Cer 18:1;O2/24:1, which increased in sEVs from WT ECs versus decreased in sEVs from CD36 −/− ECs (Fig. 5d). Lysophosphatidylcholine (LPC) also showed opposite OA-induced changes, decreasing in sEVs from WT ECs versus increasing in sEVs from CD36 −/− ECs. Thus, OA increased Cer 18:1;O2/24:1 and decreased LPC in sEVs from cells that endocytosed the caveolae with FA and the opposite occurred in sEVs from ECs where caveolae/FA endocytosis was suppressed. Effects on caveolae lipids might have contributed to the role of Src and CD36 in FA transcytosis. Src phosphorylation of Cav-1 Y14 alters Cav-1 interactions with other caveolae resident proteins, and many of these proteins bind and stabilize caveolae lipids 88 . In the case of CD36, in addition to its interaction with FA and association with Src, it might exert regulation through its binding phosphatidylserine 89 , which is enriched in caveolae and important for activation of nSMase2 and ceramide generation 68 . Additional studies are needed to examine how the FA-CD36 interaction alters caveolae lipid distribution as it relates to caveolae endocytosis and sEV secretion.
Physiological relevance of the FA-sEV pathway was shown with transwells where inhibition of the sEV pathway by GW4869 markedly suppressed FA flux from upper chamber ECs to myotubes in the lower chamber. Also upper chamber hMECs expressing the exosome marker RFP-CD63 transferred OA together with CD63 to myotubes. In addition, a mouse model with endothelial-specific expression of emGFP-CD63, click chemistry of intravenously injected alkyne OA visualized OA transfer with emGFP-CD63 to muscle fibers. Inhibition of exosome formation suppressed muscle fiber FA uptake in vivo, increased blood FAs, and reduced glucose levels but only when mice expressed CD36. Src inhibition to block Cav-1 phosphorylation also reduced FA uptake by muscle and increased blood FAs in mice. Our studies suggest that the sEV pathway plays an important physiological role in uptake of circulating FA. Additional work should further assess its quantitative contribution, particularly under different metabolic states. In this context, a previous report noted that in mice circulating exosomes have more CD36 postprandially as compared to after an overnight fast, and CD36 acquisition was important for loading BODIPY-FA into exosomes. Injection of mice with exosomes prelabeled with BODIPY-FA resulted 1.5 h later in recovery of fluorescence in the heart 90 . The origin of the postprandial CD36 expressing exosomes was not determined but might reflect in part spillover into the circulation of EC-derived FA-sEVs generated during intravascular lipolysis. How the EC FA-sEV pathway is affected by feeding, overfeeding or high fat intake, and the metabolic consequences remain to be elucidated.
The FA sEV pathway will be relevant to our understanding of how circulating FAs influence tissue crosstalk and metabolic health 4,5 and how in some cases FAs can promote metabolic disease or cancer metastasis 12,91,92 . Exosomes are critical in cell-cell communication and influence the phenotype of recipient cells by carrying microRNAs, bioactive lipids, and other regulatory factors. The FA-treated ECs generated sEVs that enhanced expression of key metabolic enzymes of both FA and glucose in myotubes. These data could help explain the reported effects of FA supply in inducing adaptations of skeletal muscle FA uptake and oxidation 93 . The sEVs also increased expression in myotubes of eNOS and that of VEGF-R2 suggesting exosomes contribute to vascular regulation and capillary maintenance. In human muscle, a 3-h bout of exercise increases gene expression of VEGF-R2, eNOS, and HK2, effects referred to as hormetic, meaning adaptive and aimed at reestablishing homeostasis 94 .
During their biogenesis, intracellular traffic, and secretion, the sEVs integrate various proteins such as membrane transporters, tetraspanins, enzymes, and proteins important for membrane fusion or for targeting sEVs to specific cells or cellular organelles, etc 95 . The cargo of the FA-sEVs likely includes proteins other than CD36 that function in FA uptake, e.g., FA acyl-CoA synthases 96 and cytosolic FA binding proteins, FABPs 97,98 . These proteins would serve as intracellular docking sites targeting the vesicles to specific cellular organelles. The FA-sEV pathway is likely to predominate in the polarized ECs. These cells control tissue FA uptake but utilize little FAs, and as we showed, transcytose the FAs for export in sEVs to parenchymal cells. However, in contrast to ECs, parenchymal cells keep most FAs for local use (e.g., cardiomyocytes, myocytes, adipocytes). In these cells, the FA-sEV which might associate with FABPs during traffic or sorting would be primarily directed to cellular organelles for processing, e.g., to lipid droplets 50 , mitochondria, or the nucleus, as highlighted in Fig. 10. This might explain the marked defect in lipid droplet formation we previously observed in the heart of mice with cardiomyocyte CD36 deletion despite presence of normal myocardial FA uptake. This observation contrasted with the findings in mice with EC CD36 deletion where uptake and lipid formation were both suppressed 9 .
Vesicular recycling of CD36 between endosomes and sarcolemma regulates cell surface CD36 and myocardial FA uptake 99 and is governed by assembly of the vacuolar H + -ATPase (V-ATPase) complex which acidifies endosomes. Endosome alkalinization pursuant to V-ATPase disassembly by FAs was reported to induce permanent CD36 relocation to the membrane with dysfunction of FA uptake 87 . Interestingly, V-ATPase also regulates the sEV pathway and a decrease in its activity promotes sEV secretion by reducing MVB acidification 69,100 . Regulation of V-ATPase in different cellular organelles could play an important role in directing the endocytosed FA vesicles. We speculate that some of the FA handling proteins, notably the FABPs will associate with the FA-EVs during their sorting and play a role in directing the vesicles in a cell type-specific manner, to lipid droplets, mitochondria, nuclei, or other sites.
The sEV FA uptake pathway might contribute to some of CD36 actions in metabolism, atherosclerosis, and immunity 11,91,101 as the findings could apply to CD36 ligands other than FAs. These could include native or oxidized lipoproteins, the multifunctional integrins, amyloid beta, pathogens 20,56 etc., and could have relevance to other CD36 actions 11,91,101 . Our findings connect FA and ceramides independent of ceramide synthesis as they show that FAs regulate membrane ceramide metabolism, which in turn regulates FA uptake. It will be important to understand the implications of this new connection in the etiology of FA-or ceramide-induced cellular pathophysiology 102 .

Methods
Mice C57BL/6 Cd36 −/− mice 27,103 and wild-type (WT) controls were used for isolation of lung microvascular cells. A mouse with EC-specific expression of emGFP tagged CD63, an exosome marker, was generated by crossing CD63-emGFP(loxP/stop/loxP) with Cdh5-Cre(ERT2) mice 104 to generate tamoxifen inducible EC-CD63(emGFP +) mice as previously described 84 . All experiments included male and female animals. The mice were maintained under standard laboratory conditions with a 12-h light/12-h dark cycle. The room temperature was maintained between 20 and 23°C, and the humidity ranged from 30 to 70%. The mice were provided with rodent chow and water throughout the study period unless indicated. Mice were euthanized using CO 2 inhalation according to institutional guidelines. Protocols were approved by the Washington University Institutional Animal Care and Use Committee and complied with ethical regulations for laboratory animal studies.

In vivo inhibition of nSMase2 and FA click chemistry
Wild-type and Cd36 −/− mice were intraperitoneally (IP) injected either with DMSO (vehicle control) or with 2.5 µg/g GW4869 105 , an inhibitor of nSMase2 for 4 days (n = 4-5/genotype/group). Src inhibitor-1 or vehicle were IP injected (2 mg/kg) for 4 days (n = 4-5/genotype/group) 106 . Mice were either studied fed (at the end of the dark period) or after a 5 h fast. Blood was collected from the tail vein and non-esterified FAs (NEFA) were measured (Wako Diagnostics, USA). For in vivo click chemistry, 5 h fasted mice (DMSO or GW4869 groups) were retroorbitally injected under 2% isoflurane 100 µl of warmed PBS with alkyne OA (to yield~25 µM OA in the circulation based on 2 mL blood volume in a 25-30 g mouse). After 30 min, mice were euthanized by cervical dislocation under 2% isoflurane and skeletal muscle processed for vapor fixation. Plantar flexor muscles were dissected from the hindlimb, frozen in liquid nitrogen cooled isopentane, and stored at −80°C until sectioning. Sections, 10 µm, were made on a cryostat (Leica Biosystems) at −20°C and mounted on pre-chilled slides, which were vapor fixed for 24 h at −20°C by inversion in pre-chilled 37% formaldehyde in a closed staining dish while keeping sections in the vapor phase. Sections were then post-fixed in the liquid phase, by inverting slides again so sections rested in the 37% formaldehyde for 30 min. Sections were then re-hydrated in PBS until staining. Confocal imaging used Zeiss LSM II Airyscan microscope.

Fatty acid uptake assay
The fatty acid uptake real-time assay 44 was performed according to the manufacturer's protocol (QBT Fatty Acid Uptake Kit, Molecular Devices). The assay was optimized for a 96-well format. Briefly, equal number of hMECs were seeded in black, clear bottom 96-well plates coated with attachment factor (AF S006100, ThermoFisher Scientific). Upon reaching confluence, cells were washed, and serum starved for 4 h. After a gentle wash with PBS the cells were treated either with vehicle control or SSO (100 µM, 30 min), GW4869 (10 µM, 10 min), Latrunculin B (2.5 µM, 30 min), Src inhibitor-1 (10 µM, 2 h), or ROCK inhibitor (10 µM, 30 min). The cells were washed with HBSS, 100 µL HBSS was then added, and basal fluorescence measured with a prewarmed (37°C) microplate reader (bottom read, excitation 488/ emission 515, filter cutoff 495 nm, 21 s read). With a multichannel pipette, HBSS was removed, 100 µL warmed QBT (protected from light) added, and fluorescence measured. Background reading with HBSS were subtracted from QBT readings to get final fluorescence. Data were plotted as the calculated area under the curve (AUC).

Transmission electron microscopy (TEM)
Subcellular visualization of palmitic acid in endothelial cells used palmitoyl nanogold (Nanoprobes, Yaphank, NY) for 10 min before fixing cells with 4% PFA (Polysciences Inc., Warrington, PA) in 100 mM PIPES/ 0.5 mM MgCl 2 , pH 7.2 for 1 h at 4°C. Samples embedded in 10% gelatin were incubated overnight at 4°C in 2.3 M sucrose/20% polyvinyl pyrrolidone in PIPES/MgCl 2 , then trimmed, frozen in liquid nitrogen and sectioned (Ultracut UCT7 cryo-ultramicrotome Leica Microsystems Inc., Bannockburn, IL). Ultrathin sections (50 nm) placed on nickel grids freshly coated with glow discharged carbon, were washed in PIPES buffer and incubated with GoldEnhance EM Plus (Nanoprobes) per the manufacturer's instructions. Development times determined 4 min as optimal for gold deposition onto nanogold particles and minimal background. Grids were rinsed extensively in deionized water, stained with 0.3% uranyl acetate/2% methyl cellulose, and viewed on a JEOL 1200 EX transmission electron microscope (JEOL USA Inc., Peabody, MA) with an AMT 8-megapixel digital camera and AMT Image Capture Engine V602 (Advanced Microscopy Techniques, Woburn, MA).

Isolation and characterization of sEV
EC conditioned media were centrifuged to remove cell debris (2000 × g, 20min) and microvesicles (15,000× g, 30 min). Postmicrovesicle supernatant was incubated with ExoQuick TC (Systems Biosciences) overnight at 4°C and centrifuged (1500 × g, 30 min) to pellet sEVs. The pellets were resuspended with PBS for Nanoparticle Tracking Analysis (Nanosight NS300, Malvern Pananalytical) or in 1X RIPA buffer for western blot and marker analysis. The sEVs from cells incubated with nanogold palmitic acid were subjected to TEM as above.
Isolation and characterization of 3 H-oleate loaded sEV mMECs and hMECS incubated with 3 H-oleate solution complexed with BSA. sEVs were isolated as described above. sEV were dissolved in Article chloroform/methanol (1:1, by vol) and lipids were separated by thin layer chromatography using petroleum ether/diethyl ether/acetic acid (70:30:1, by vol). Lipid classes were visualized by exposure to iodine vapor, the corresponding bands scraped, and associated radioactivity determined by liquid scintillation counting.

Uptake of EC-generated FA-sEVs by myotubes in transwells
For visualizing sEV transfer from ECs to myotubes, hMECs were transduced with RFP-CD63 lentiviral particles (Systems Bioscience). After 7 days of antibiotic selection the cells were grown on collagencoated transwells (Falcon, 0.4 µm pore) in 24-well plates until 3-5 days post confluence, assessed by trans-endothelial electric resistance (TEER) 80  Human skeletal muscle myoblasts (hSMM) on coverslips in skeletal muscle growth media (Lonza) were grown and differentiated into myotubes 55 below the transwells. Serum-starved (4 h) hMECs were treated with alkyne OA (25 µM) added to upper chamber and bottom myotubes were collected 15 or 30 min later for immunofluorescence (Zeiss LSM 880 Airyscan Confocal Microscope, 40X or 63X). Images were processed using Zeiss Zen (Blue) Imaging. ImageJ was used to determine mean intensity.
RNASeq on control and CD36-knockdown hMEC hMECs were treated with either control or CD36 siRNA as above. 72 h after treatment, total RNA was isolated using TRIzol Reagent (Ther-moFisher Scientific) followed by column-based sample cleanup (RNeasy, Qiagen). Samples were submitted to the Washington University Genome Technology Access Center (GTAC) for library preparation, sequencing and read alignment. Normalization and differential gene expression was determined using the integrated Differential Expression and Pathway Analysis (iDEP) web platform 108 .

Preparation and LC/MS analysis of sEV lipids
Samples were collected and aliquots representing the same EV counts were dried under nitrogen and resuspended in 2:2:1 acetonitrile:methanol:H 2 O (v/v) (for polar metabolites, e.g., fatty acids, phospholipids) or in 100% isopropanol (for less-polar metabolites, e.g., ceramides). The samples were mixed on an orbital shaker (360 rpm) for 1 min at room temperature before the LC/MS analysis. LC/MS analysis was performed on an Agilent 1290 Infinity II LCsystem coupled to an Agilent 6545 QTOF mass spectrometer with a dual Agilent Jet Stream electrospray ionization source. Analysis of ceramides and fatty acids was further verified in high resolution using an Orbitrap ID-X Tribrid mass spectrometer (Thermo Scientific). A Vanquish Horizon UHPLC system, was interfaced with the mass spectrometer via electrospray ionization in both positive and negative ion mode with a spray voltage of 3.5 and 2.8 kV, respectively. Lipids were separated on a CORTECS UPLC C18 column (2.1 × 100 mm, 1.6 µm; part No. 186007095) including a Waters UPLC HSS VanGuard Pre-Column (2.1 × 5 mm, 1.8 µm; part No. 186007949) at a temperature of 60°C and a flow rate of 250 mL/min. The mobile phases consisted of A: 60% acetonitrile, 40% water, 0.1% formic acid, 10 mM ammonium formate, 2.5 mM medronic acid, and B: 90% 2-propanol, 10% acetonitrile, 0.1% formic acid, 10 mM ammonium formate (in 1 mL water). The following linear gradient was used: 0-2 min, 30% B; 17 min, 75% B; 20 min, 85% B; 23-26 min, 100% B; 26 min, 30% B followed by a re-equilibration phase of 5 min. Lipids were detected in positive ion mode with following source parameters: gas temperature 250°C, drying gas flow 11 L/min, nebulizer pressure 35 psi, sheath gas temperature 300°C, sheath gas flow 12 L/min, VCap 3000 V, nozzle voltage 500 V, Fragmentor 160 V, Skimmer 65 V, Oct 1 RF Vpp 750 V, and m/z range 50-1700. Data were acquired under continuous reference mass correction at m/z 121.0509 and 922.0890 in positive ion mode. Samples were randomized before analysis and a quality-control (QC) sample was injected to monitor instrument signal stability.
Acquiring MS/MS data, data processing, and normalization MS/MS spectra for lipids were acquired using an iterative approach in the MassHunter Acquisition Software (Version 10.1.48, Agilent Technologies) on an Agilent 6545 QTOF. Source settings for MS1 data acquisition were used. MS/MS spectra were acquired at a scan rate of 3 spectra/s with different intensity thresholds and collision energies of 10, 20, and 40 V to increase identification rates. For the ID-X Orbitrap, data were acquired in data-dependent acquisition (DDA) by using the built-in deep scan option (AcquireX) with a mass range of 67-900 m/z. The MS/MS scans were acquired at 15 K resolution and cross referenced to a library generated from NIST SRM 1950 plasma sample in both positive and negative ion mode with different collision energies in the range of 20 NCE to 50 NCE for HCD and 30 NCE for CID to maximize identifications.
Lipid iterative MS/MS data were annotated with the Agilent Lipid Annotator software. All data files were then analyzed in Skyline-daily (Version 22.2.1.256) to obtain peak areas, m/z values of the metabolite and lipid target lists, obtained from the metabolite identification workflow, which had at least an MS/MS match to an online library, were extracted under consideration of retention times.

Statistical analyses
All data presented are representative of at least three independent experiments, unless indicated otherwise. Quantifications are shown as means ± standard errors (S.E.). Statistical significance, p value was evaluated using unpaired Student's t test when comparing between two groups. For comparison between more than two groups one-way ANOVA or two-way ANOVA analysis was used followed by post hoc Tukey for multiple comparisons as indicated (****P ≤ 0.0001, ***P ≤ 0.001, **P ≤ 0.01, *P ≤ 0.05, ns P > 0.05).

Reporting summary
Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability
Source data files, including uncropped blots, are provided with this paper (Source data file). Lipidomic data generated in this study is deposited at metabolomics workbench and can be accessed at https:// doi.org/10.21228/M8PH89. RNA-seq data in this study have been deposited in the Gene Expression Omnibus database under accession code GSE235988. All other relevant data could be obtained from the corresponding authors Vivek Peche and Nada Abumrad upon request. Source data are provided with this paper.
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